Caterpillar–parasitoid interactions: species-specific influences on host microbiome composition

Abstract There is increasing evidence that host–parasitoid interactions can have a pronounced impact on the microbiome of host insects, but it is unclear to what extent this is caused by the host and/or parasitoid. Here, we compared the internal and external microbiome of caterpillars of Pieris brassicae and Pieris rapae parasitized by Cotesia glomerata or Cotesia rubecula with nonparasitized caterpillars. Additionally, we investigated the internal and external microbiome of the parasitoid larvae. Both internal and external bacterial densities were significantly higher for P. brassicae than P. rapae, while no differences were found between parasitized and nonparasitized caterpillars. In contrast, parasitism significantly affected the composition of the internal and external microbiome of the caterpillars and the parasitoid larvae, but the effects were dependent on the host and parasitoid species. Irrespective of host species, a Wolbachia species was exclusively found inside caterpillars parasitized by C. glomerata, as well as in the corresponding developing parasitoid larvae. Similarly, a Nosema species was abundantly present inside parasitized caterpillars and the parasitoid larvae, but this was independent of the host and the parasitoid species. We conclude that parasitism has pronounced effects on host microbiomes, but the effects depend on both the host and parasitoid species.


Introduction
Most insects harbour a variety of micr oor ganisms whose div ersity and roles are only recently being better understood (Engel and Moran 2013, Douglas 2015, Muñoz-Benavent et al. 2021 ).Their internal microbiomes either comprise a stable assemblage of micr oor ganisms that can be consistently detected in larval and adult hosts (Bright andBulgheresi 2010 , Engel andMoran 2013 ) or harbour transient gut microbes (Hammer et al. 2017(Hammer et al. , 2019 ) ).These micr oor ganisms ma y pla y important roles in insect behaviour, food digestion, nutrition, detoxification, and protection of their host against abiotic stress , pathogens , and parasites (Douglas 2015 ).Similarly, the external surfaces of insects (i.e. the exoskeleton) are commonly inhabited by microorganisms.Unlike the internal microbiome, the external insect microbiome is often composed of a diverse group of nonspecialized environmental microorganisms that v ary significantl y with geogr a phic location and habitat (Park et al. 2019 ).
The composition and diversity of insect micr obiomes ar e affected by a wide range of factors, including host phylogeny, life stage, diet, and habitat (Behar et al. 2008, Ottesen and Leadbetter 2010, Yun et al. 2014, Chen et al. 2016, Shao et al. 2024 ).Additionall y, ther e is incr easing e vidence that the microbial commu-nity composition and diversity in insects is str ongl y influenced by host-par asite inter actions (but see Liu et al. 2020 ).P ar asites like helminths and protozoa residing in the insect gut may alter the composition of the gut micr obiome (Fr edensbor g et al. 2020 ).Similarl y, insect-par asitic nematodes (Vicente et al. 2016 ) and koinobiont parasitic wasps (parasitoids) have been shown to modify the internal microbiome of their hosts (P olenogo va et al. 2019 , Cavic hiolli de Oliv eir a and Consoli 2020 , Gao et al. 2021, Gloder et al. 2021, Zhang et al. 2022, Wang et al. 2023, Gw ok y aly a et al. 2024 ).
Koinobiont parasitoids are important secondary consumers in arthr opod comm unities and k e y natur al enemies of a gricultur al pests .T he y de posit their eggs inside or outside their hosts, and their larv ae par asitize the hosts while k ee ping them ali ve for a certain period of time (Schafellner et al. 2004 ).The parasitoid larvae can alter host behaviour such as food preference (Smilanich et al. 2011 ) and food intake and utilization (Rossi et al. 2014 ), which in turn may impact the diversity and composition of the host insect microbiomes (Yun et al. 2014 ).Furthermor e, adult par asitoids may transfer some of their microbiota during oviposition and alter the microbiome of their host both directly and indirectly (Douglas 2015 , Gloder et al. 2021, Gw ok y aly a et al. 2024 ).Research has demonstr ated that par asitoid symbionts and v enom injected with the wasp eggs can manipulate host physiology and suppress the host immune system to benefit the survival of the parasitoid's offspring (Strand and Pech 1995 ).At the same time, this process may also affect the regulation of gut micr obes, ther eby indir ectl y changing the host microbiome (Cavicchiolli de Oliveira and Consoli 2020 ).
Ther efor e, we hypothesize that the internal microbiomes of par asitized insects ar e to a lar ge extent determined by c har acteristics of both the host and parasitoid species.Similarly, we predict that the microbiome of the parasitoid larvae that develop in the host is determined by features of both the host and parasitoid species .Con versely, given that parasitoids parasitize the interior of their hosts, we expect that they do not, or to a lesser extent, affect the external microbiome of their host (Gloder et al. 2021, Bourne et al. 2023 ).To test these hypotheses, we compared both the internal and external microbiomes of parasitized and nonparasitized hosts and examined whether differ ences wer e mainl y driv en by the host, the parasitoid, or a combination of both.Furthermore, we asked which microbes wer e commonl y tr ansferr ed to the hosts thr ough par asitism.We also assessed the microbiomes of the de v eloping par asitoid larv ae and inv estigated to whic h extent they ar e influenced by the host, the parasitoid or their interaction.To this end, we used the lar ge cabba ge white Pieris brassicae and the small cabbage white Pieris rapae (Lepidoptera: Pieridae) and their main koinobiont endoparasitoids Cotesia glomerata and Cotesia rubecula (Hymenoptera: Braconidae) as study species.Previous research using P. brassicae and C. glomerata has shown that parasitism by C. glomerata has a major impact on the host microbiome (Gloder et al. 2021, Bourne et al. 2023 ).Ho w e v er, the specific contributions of the host and parasitoid species to these alterations remain to be fully elucidated.

Study species
Pieris rapae has a natur al r ange acr oss Eur ope, North Africa, and Asia, but has also been found in North America, Australia, and New Zealand.In contrast, P. brassicae is less widely distributed and mainly occurs in Europe, Asia, and North Africa.Both species are important pests on many crop species belonging to the family Br assicaceae suc h as cabba ge, cauliflo w er, Brussels sprouts, and r a pe .Pieris brassicae la ys eggs in clusters of 10-100 eggs whereas P. rapae lays single eggs, leading to gregarious and solitary larvae, r espectiv el y (Davies and Gilbert 1985 ). Cotesia glomerata is a gregarious koinobiont wasp that parasitizes a wide range of caterpillars of pierid butterflies, but P. brassicae and P. rapae are its main hosts (Brodeur et al. 1996 ).On a verage , adult females of C. glomerata lay around 20 eggs in a host caterpillar per oviposition e v ent (Br odeur et al. 1996 ).In contrast, C. rubecula is a solitary parasitoid and has long been considered to be specific to P. rapae (Shenefelt 1972 ), but it may also parasitize P. brassicae larvae (Brodeur et al. 1996(Brodeur et al. , 1998 ) ).
Once the egg(s) hatch, the larvae of both parasitoid species feed on the cater pillar's haemol ymph while the cater pillars ar e still aliv e. Larvae of C. glomerata emerge from their caterpillar host ∼15-20 days after parasitization, while it takes around 10-15 days for C. rubecula larvae to emerge and pupate outside of the host.At that time cater pillars ar e gener all y in the last instar (L5) when parasitized by C. glomerata , while they are in the late third (L3) instar for C. rubecula .This process eventually kills the caterpillar host (Brodeur et al. 1996 ).

Experimental set-up
The insects used in this study were taken fr om lab-r ear ed populations that were originally collected from agricultural fields in the surr ounding of Wa geningen Univ ersity & Researc h, the Netherlands.Both Pieris species were reared and maintained on Brussels sprouts plants ( Brassica oleracea L. var.gemmifera ) in separate cages in a greenhouse compartment (21 ± 1 • C, 25%-35% RH, 16:8 h light/dark).Male and female butterflies w ere allo w ed to freely mate in the cage and lay their eggs on the plants.Adult butterflies were fed with a saturated sugar solution.Cotesia glomerata and C. rubecula wer e r ear ed in individual cages in distinct greenhouse compartments under the same conditions, utilizing P. brassicae caterpillars as hosts for both species.When the parasitoid larvae had pupated, pupae were collected and transferred to a smaller cage without plants.Emerged parasitoids were provided with 10% honey-water solution until they were used in the experiments.
When P. brassicae and P. rapae larvae had hatched, first instar larvae originating from the same egg-batch were collected from our r earing, separ ated in groups of similar size ( c .20 individuals) and subjected to thr ee tr eatments: (1) par asitization by C. glomerata , (2) parasitization by C. rubecula , or (3) untreated (control caterpillars).Eac h cater pillar was individuall y par asitized as described in Cun y et al. ( 2022 ).In brief, cater pillars wer e consider ed par asitized when the parasitoid female had inserted its ovipositor in the caterpillars for at least 5 s for C. glomerata or 1 s for C. rubecula .Next, caterpillars fr om eac h combination of host and parasitoid species, as well as untr eated cater pillars, wer e placed in separ ate ca ges on wild cabbage plants ( B. oleracea , grown from seeds from Kimmeridge, UK; Gols et al. 2008 ) in the same greenhouse compartment (21 ± 1 • C, 25%-35% RH, 16:8 h light/dark), until the caterpillars were used for microbiome sampling.

Microbiome sampling
When par asitoid larv ae wer e close to egr ession, eight cater pillars fr om eac h tr eatment wer e r andoml y pic ked fr om their r espectiv e ca ge for micr obiome sampling (48 cater pillars in total; 8 × 2 caterpillar species × 3 treatments).At that time, caterpillars parasitized by C. glomerata were in the early fifth instar stage, while cater pillars par asitized b y C. rubecula w ere in the late thir d instar stage.Cotesia rubecula is known to arrest host development at the third instar stage (Harvey et al. 1999 ), while C. glomerata allows its host to r eac h the final instar sta ge (Harv ey et al. 2012 ).Nonparasitized P. brassicae and P. rapae cater pillars wer e in the early fifth instar sta ge.Pr eliminary anal ysis of a small number of P. brassicae caterpillars sho w ed no significant variation in microbiome composition among the final instar sta ges.Cater pillars wer e collected using sterilized tweezers treated with 70% ethanol.Additionally, glov es wer e worn that were also sterilized with 70% ethanol before a caterpillar was sampled.Each caterpillar was put individually in a plastic sterile container (12 cm diameter; 5 cm height) containing tissue paper (to absorb frass and moisture) with a pierced lid.Cater pillars wer e starv ed ov ernight at r oom temper atur e in the same containers to allow the insects to empty their gut content.Subsequently, both the external (cuticle associated) and internal microbiome of the caterpillars and parasitoid larvae were sampled as described in Gloder et al. ( 2021 ) ( Supplementary Table S1 ).
Briefly, the external microbiota of the caterpillars were collected by putting each caterpillar in a 2-ml microcentrifuge tube containing 1 ml of phosphate-buffered saline with 0.01% Tween80 (PBS-T), and vortexing it for 20 s .T his washing solution was then used as a sample for the caterpillar's external micr obiome.Next, cater pillars wer e surface-sterilized with sodium hypochlorite (2.5%) and washed again two times in PBS-T (Gloder et al. 2021 ), and then dissected in the proximity of a Bunsen burner to obtain internal host and par asitoid larv ae samples; Supplementary Fig. S1 ).Caterpillars were pinned onto a sterile dissection dish with flame-sterilized needles and cut open along the entire length of the caterpillar.Parasitoid larvae were collected with a sterilized pair of tweezers and put in a clean microcentrifuge tube.When necessary, some drops of sterile water were applied on top of the dissected caterpillars in order to ease the collection of the parasitoid larvae and to ensure that all larvae were r etrie v ed; Supplementary Fig. S1 ).When caterpillars were parasitized by C. glomerata all the parasitoid larvae present in a single host were pooled and treated as a single sample.To avoid contamination of the parasitoid larvae with host microbes, the dissection was performed very carefully, aiming to not damage the host gut or any other tissues other than the host cuticle.Furthermor e, dissection dishes wer e cleaned after eac h dissection, first with sodium hypochlorite (2.5%), then with ethanol (70%), and finally flooded with sterile water followed by air drying in sterile conditions .On a v er a ge, 22.7 C. glomerata larvae ( c . 2 mm in size) wer e r ecov er ed fr om P. rapae cater pillars (r ange: 8-37; median: 23), while 24.6 C. glomerata larvae were retrieved from P. brassicae caterpillars (range: 18-37; median 24) .When caterpillars were parasitized by C. rubecula , in e v ery host a single par asitoid larv a was found (3-4 mm).The rest of the body of the caterpillars was then homogenized as described before (Gloder et al. 2021 ) to r epr esent the internal host microbiome .T herefore , the remaining portion of each caterpillar was placed in a 2-ml tube containing a mixture of glass beads (three beads of 2 mm and two beads of 5 mm in diameter) and 1 ml PBS-T.The samples were then subjected to two consecutive cycles of 10 s at a speed of 5.5 m/s in a Bead Ruptor Elite (Omni international, Kennesaw, USA).The external and internal microbiome of the recovered parasitoid larvae were also sampled separ atel y following the same protocol, but with a smaller working volume of PBS-T (500 μl instead of 1 ml).

DN A extr action and molecular anal ysis
Genomic DN A w as isolated from all external and internal samples (500 μl) using the Po w erPr o Soil Kit (Qia gen, Hilden, Germany) following the manufacturer's instructions, with one modification: in the second step of the protocol the use of a vortex adapter was replaced by two cycles of 30 s (with a 10 s break in between) in the Bead Ruptor Elite at a speed of 5.5 m/s.Two negative controls in which the sample material was replaced by sterile, DNA-free w ater w as included to confirm the absence of r ea gent contamination.DN A samples w er e then subjected to molecular anal ysis.First, bacterial presence and density was assessed by a qPCR (quantitativ e r eal-time Pol ymer ase Chain Reaction) assay using the universal primers 515F and 806R (Ca por aso et al. 2011 ), amplifying the V4 region of the bacterial 16S ribosomal RN A (rRN A) gene, as described pr e viousl y (Gloder et al. 2021 ).Briefly, qPCR amplification was performed using the StepOnePlus™ RealTime PCR (Pol ymer ase Chain Reaction) System (Applied Biosystems, Foster City, CA, USA).Eac h r eaction mixtur e contained 0.2 μl of each primer (20 μM), 10 μl of the iTaq Universal SYBRGreen supermix (Biorad, Hercules, CA, USA), 8.6 μl of sterile distilled water, and 1 μl of template DNA.The thermal cycling protocol consisted of an initial denaturation step at 95 • C for 2 min follo w ed b y 40 amplification cycles of 15 s at 95 • C and 1 min at 60 • C. Fluorescence (520 nm) was measured at the end of the elongation phase in each cycle .For each sample , the threshold c ycle (C T ) w as calculated using StepOne™ software, and the baseline was set automatically abov e an y noise.All qPCR r eactions wer e performed in duplicate, and each run included a negative control where template DNA was replaced with sterile water.Additionally, a 10-fold dilution series of the targeted DNA fragment (ranging from 1 ng/ μl to 1 fg/ μl, measured with a Qubit fluorometer; Invitrogen, Carlsbad, USA) was included in each run to establish a calibration curve for calculating the number of gene copy numbers per μl DNA extract in the investigated samples (Lee et al. 2006 ).This dilution series w as obtained b y first amplifying the V4 region of a r efer ence str ain ( Pseudomonas sp.ST09.08/02) using the primers 515F and 806R, and diluting it.The detection limit of the assay was set at a C T value of 34, whic h corr esponded to the lo w est C T value obtained for one of the blanks.Results of the gene copy numbers from the qPCR amplification are shown in Supplementary Table S2 .
Additionall y, for eac h sample the V4 r egion was amplified using Illumina barcoded versions of the same primers to assess the diversity and composition of the bacterial communities in the samples.Primers were designed according to Kozich et al. ( 2013 ) (dual index sequencing strategy) ( Supplementary Table S3 ).In addition to the different DNA samples, three negative PCR controls (in which DNA template was replaced by DNA-free water) were included, as well as a DNA mock community sample that was composed of a number of bacterial species that likely occur in or on insects (Gloder et al. 2021 ) ( Supplementary Table S4 ).PCR amplification, libr ary pr epar ation, sequencing, and bioinformatics analysis were performed as described pr e viousl y (Gloder et al. 2021 ).Briefly, amplification was performed in a reaction volume of 40 μl, consisting of 2 μl DNA, 0.5 μM of each primer, 150 μM of each dNTP, 1 × Titanium Taq PCR buffer and 1 × Titanium Taq DNA pol ymer ase (Takar a Bio, Saint-Germain-en-La ye , France) with the follo wing c ycling protocol: 94 • C for 120 s, follo w ed b y 35 c ycles of 45 s at 95 • C, 45 s at 59 • C, and 45 s at 72 • C, and a final elongation step of 10 min at 72 • C. Amplicons from all insect samples and contr ols wer e purified using Agencourt AMPur e XP ma gnetic beads (Beckman Coulter Genomics GmbH, South Plainfield, UK) following the manufacturer's instructions.Subsequently, a Qubit high sensitivity fluorometer (Invitrogen) was used to measure the concentration of the purified amplicons, and each sample was pooled at equimolar concentrations.After ethanol precipitation, the amplicon library was loaded onto a 1.5% agarose gel, and the target band was excised and purified using a QIAquick Gel Extraction Kit (Qia gen).Following gel extr action, the concentr ation of the libr ary was measured again, diluted to 2 nM, and then sent for sequencing at the Centre for Medical Genetics of the University of Antwerp (Antwerp, Belgium) using an Illumina MiSeq sequencer with a v2 500-cycle r ea gent kit (Illumina, San Diego, USA).
Bacterial sequences were received as demultiplexed FASTQ files, with barcodes and primer sequences r emov ed.P air ed-end r eads wer e mer ged using USEARCH (v11.0.667) to gener ate consensus sequences (Edgar 2013 ), with no more than 10 mismatches allo w ed in the ov erla p r egion.Subsequentl y, r eads shorter than 190 bp or with a total expected error threshold above 0.05 were discar ded.Sequences w ere then classified into zer o-r adius oper ational taxonomic units (zOTUs; Edgar 2016 ), also known as amplicon sequence variants (Callahan et al. 2017 ) by the UNOISE3 algorithm as implemented in USEARCH (Edgar and Fl yvbjer g 2015 ).
The obtained dataset was decontaminated in R (v3.5.2) (R Core Team 2018 ) using microDecon (v1.0.2) (McKnight et al. 2019 ) to r emov e contaminants based on zOTU pr e v alence in the insect samples versus the mean of the three PCR controls (Davis et al. 2018 ).At the same time, the DNA extr action contr ols wer e r emov ed fr om the dataset since they yielded onl y v ery low sequence numbers and no additional zOTUs in comparison with the PCR controls.Also, no band was obtained for the DNA extraction controls when loading the samples on an a gar ose gel, indicating that the DNA extraction kits were free of bacterial contamination.Subsequently, zOTUs occurring below a 0.1% relativ e abundance thr eshold in a giv en sample wer e discarded in that sample prior to further analysis (Gloder et al. 2021, Gorrens et al. 2022, Ijdema et al. 2022 ).In this way, analysis of the moc k comm unity onl y yielded the expected comm unity members ( Supplementary Table S5 ), demonstrating the robustness of our method.Finally, to correct for uneven sequence numbers, the number of sequences was r ar efied to 2000 sequences per sample, while samples with less sequences w ere discar ded from the analysis .T he taxonomic origin of each zOTU was determined with the SINTAX algorithm as implemented in USEARCH based on the SILV A Living T r ee Pr oject v123.The identity of the most important zOTUs was also verified with a BLAST search in GenBank against type materials.When no significant similarity values were found ( < 97% identity), the BLAST analysis was performed against the entir e database.Ov er all, r esults obtained by the BLAST anal ysis matc hed v ery well with those obtained with the SINTAX algorithm in USEARCH ( Supplementary Table S5 ).

Da ta anal ysis
Data analysis was performed on distinct datasets, one comprising samples from the caterpillars and another with samples from the par asitoid larv ae.Additionall y, the samples wer e categorized into internal and external samples.To test whether bacterial densities (determined by qPCR), expressed as the number of 16S rRNA gene copies per μl DN A extract, w ere affected b y caterpillar host species ( P. brassicae or P. rapae ), parasitism status (parasitized by C. glomerata , parasitized by C. rubecula , or nonparasitized) and their inter action, a Sc heir er-Ray-Har e test in rcompanion pac ka ge in R (Mangiafico 2023 ) was performed for both the internal and external caterpillar samples (test performed on logarithmic values).This test is a nonparametric test used for a tw o-w ay factorial design (data did not meet the assumption of equal variances, as assessed with a Le v ene test).The same test was performed on samples collected from the parasitoid larvae residing within the parasitized caterpillars.For statistical analysis, samples in which bacteria could not be detected using qPCR but were detected through sequencing, were assigned to the qPCR detection threshold of 2.95 × 10 2 16S rRNA gene copies per μl DNA extr act, whic h is equiv alent to a C T value of 34.
To assess whether the depth of our sequencing a ppr oac h was sufficient to ca ptur e the bacterial div ersity in the samples, r arefaction curves ( Supplementary Fig. S2 ) were generated after rarefying the data to 2000 sequences per sample using the Phyloseq pac ka ge in R (McMurdie and Holmes 2013 , R Core Team 2018 ).The same pac ka ge was used to determine zOTU richness (i.e. the number of observed zOTUs) and Shannon diversity for each sample.A tw o-w ay analysis of variance (ANOVA) was used to assess whether host cater pillar species, par asitism status, and their inter action affected zOTU richness and Shannon diversity in the caterpillar samples .T he same analysis was performed to assess whether host caterpillar and parasitoid species, and their interaction, affected zOTU richness and Shannon diversity in samples from the par asitoid larv ae.Bacterial comm unity composition was visualized using nonmetric multidimensional scaling (NMDS) with the Bray-Curtis coefficient as distance measure in the R package vegan, based on r elativ e abundance data.To test the hypothesis that cater pillar bacterial comm unities differ ed between host species and parasitism status, permutational analysis of variance (PER-MANOVA) was performed on the same data set using the 'adonis' function in the vegan package (Oksanen et al. 2015 ).Host species, parasitism status, and their interaction were included as fixed factors in the anal ysis.Similarl y, PERMANOVA was performed on the parasitoid larvae data to assess whether bacterial community composition within and on the parasitoid larvae differed between host caterpillars and parasitoid species, and whether there was an interaction effect.Statistical significance was tested using 1000 permutations .T his analysis and the NMDS visualization were repeated on a reduced dataset where zOTUs belonging to the same famil y wer e mer ged into famil y-le v el phylotypes .T he sequence data obtained in this study has been submitted in the Sequence Read Arc hiv e at NCBI under Biopr oject PRJNA1082293.
Indicator species analyses using the R pac ka ge 'indicspecies' were performed to investigate whether zOTUs could be assigned to specific tr eatments.Anal yses wer e performed separ atel y for eac h cater pillar species and for external and internal microbiomes.A complementary co-occurrence matrix was calculated and visualized using the 'co-occur' R pac ka ge (Griffith et al. 2016 ) using the same datasets.Finally, Kruskal-Wallis tests were used to assess whether the r elativ e abundance of individual zOTUs differ ed significantl y among tr eatments.Anal yses wer e r estricted to the 21 most abundant zO TUs , occurring at a mean r elativ e abundance > 1% in at least one of the caterpillar treatment groups.

Bacterial density
Absolute bacterial densities (calculated b y qPCR) w ere significantly higher for caterpillars of P. brassicae than for those of P. rapae , both externally and internally, and parasitism status did not impact this result (Table 1 ; Fig. 1 A and B).Larvae from both parasitoid species had a higher external bacterial density when infecting P. brassicae than when infecting P. rapae .This difference was mor e pr onounced in C. glomerata larv ae than in C. rubecula larv ae (Table 1 ; Fig. 1 C).For the internal samples of the parasitoid larvae, regardless of host species, there was a slightly, but significantly higher internal bacterial density in larv ae fr om C. rubecula than in larv ae fr om C. glomerata (Table 1 ; Fig. 1 D).

Bacterial di v ersity and comm unity composition
After quality filtering, r emov al of potential contaminants and rarefying to 2000 sequences per sample, a total of 658 zOTUs were retained in the analysis ( Supplementary Table S5 ), covering a total of 144 samples ( Supplementary Table S1 ).In gener al, r ar efaction curv es a ppr oac hed satur ation ( Supplementary Fig. S2 ), indicating that our sequencing depth of 2000 reads per sample was sufficient to cover the bacterial diversity in the samples.Tw o-w ay ANOVA of the caterpillar external microbiomes revealed no significant differences in zOTU richness between the two host caterpillars (Fig. 2 A), while a significant difference was found in Shannon diversity (Fig. 2 B; Table 2 ).This indicates that while the number of bacterial species is similar, the distribution and abundance of those species differ between the caterpillar hosts.
Although parasitism did not significantly affect zOTU richness or Shannon diversity in the external caterpillar samples, P. rapae caterpillars parasitized with C. rubecula sho w ed a higher bacterial richness and diversity (Table 2 ; Fig. 2 A and B).The internal microbiomes sho w ed significant differences betw een the tw o caterpillar species, both in terms of zOTU richness and Shannon  di versity.Higher n umbers of bacterial zOTUs and gr eater div ersity were found in P. rapae than in P. brassicae (Table 2 ; Fig. 2 C and D).Furthermor e, par asitism had a significant effect on the internal cater pillar micr obiomes, with a mor e pr onounced effect in P. rapae than in P. brassicae , both for richness and diversity.Nonparasitized P. brassicae contained an av er a ge of 1.4 (range 1-2) zOTUs, whic h incr eased to 3.3 (r ange 3-4) when par asitized by C. glomerata and to 2.4 (range 2-4) when parasitized by C. rubecula .In contrast, uninfected P. rapae caterpillars harboured an average of 35.1 (range 9-64) zO TUs , while this was only 2.1 (range 2-3) and 11.3 (3-46) when parasitized with C. glomerata and C. rubecula , respectiv el y (Table 2 ; Fig. 2 C and D).P ar asitoid larv ae had a higher zOTU richness and Shannon diversity in both the external and internal samples of C. rubecula compared to C. glomerata , and this difference in diversity was more pronounced when parasitizing P. rapae than when parasitizing P. brassicae (Table 2 ; Fig. 2 E-H).PERMANOV A analyses (T able 3 ; Supplementary Table S6 ) sho w ed significant differences in both the external and internal bacterial community composition between caterpillars of P. brassicae and P. rapae , as w ell as betw een the differ ent tr eatments (Table 3 ; Fig. 3 A and B).Ho w e v er, the effect of parasitism was mor e pr onounced in samples fr om P. rapae compar ed to P. brassicae (Table 3 ; Fig. 3 A and B).The external microbiome of para-sitoid larvae also differed significantly between both parasitoid species and between larvae collected from P. brassicae and P. rapae (Table 3 ; Fig. 3 C).Mor eov er, the inter action between host species and parasitoid species was statistically significant for the external parasitoid samples.In contrast, there was a significant difference between the internal microbiome of larvae of the two parasitoid species (Table 3 ; Fig. 3 D), while no significant differences were found between host species, nor was there a significant interaction effect (Table 3 ; Supplementary Table S6 ).When repeating the analysis at the family level, the same patterns were observed ( Supplementary Fig. S3 ; Supplementary Table S7 ).

Taxonomic classification, incidence, and relati v e abundance of caterpillar-host microbes
Bacteria found on and inside the anal ysed cater pillars r epr esented se v er al envir onmental and insect-associated species belonging to diverse phyla, with the most abundant species belonging to Pseudomonadota (Proteobacteria), Bacillota (Firmicutes), and Actinomycetota (Actinobacteria) ( Supplementary Table S5 ).In gener al, cater pillar micr obiomes wer e dominated by a limited number of bacterial species (Fig. 4 ; Supplementary Fig. S4 ).In particular, irr espectiv e of par asitism status, both the external and F igure 2. Boxplots sho wing alpha div ersity (zOTU ric hness and Shannon index) comparisons of the external and internal micr obiomes of the differ ent (A-D) caterpillars and (E-H) parasitoid larvae samples studied.Pieris brassicae and P. rapae caterpillars were parasitized with either C. glomerata (CG) or C. rubecula (CR), or r emained unpar asitized (UN).The lo w er and upper whiskers corr espond to the minim um and maxim um v alues, with the bar in the middle marking the median value while dots represent outliers.internal microbiomes of P. brassicae caterpillars were dominated by a single zO TU (zO TU1), identified as Enterococcus sp.This bacterium was found at an av er a ge r elativ e abundance of 84.1% and 90.2% on and inside P. brassicae cater pillars, r espectiv el y, while it was less abundant on (10.2%) and inside (6.4%) P. rapae caterpillars .Moreo ver, the bacterium was present in all analysed P. brassicae samples, but was absent in any P. rapae caterpillar sample parasitized by C. glomerata (Fig. 4 ).In the external microbiome, Enterococcus sp. was found in five out of six nonparasitized P. rapae caterpillars and in five out of seven caterpillars parasitized by C. rubecula , while in the internal microbiome it was found in six out of se v en nonpar asitized individuals and in fiv e out of se v en individuals parasitized by C. rubecula (Fig. 4 ).
A few bacterial species were common and abundant on or inside nonparasitized P. rapae caterpillars, while they were rare or absent on or inside P. rapae caterpillars that were parasitized.In the external microbiome , zO TU5, identified as Serratia sp., was consistentl y pr esent on all nonpar asitized P. rapae cater pillars with an av er a ge r elativ e abundance of 56.2%, while it was detected on only a few parasitized individuals at a lo w er r elativ e abundance .Additionally, zO TU7, an unidentified member of the Enter obacteriaceae famil y, and zOTU6, identified as Pseudomonas sp., wer e both pr esent in all internal samples fr om nonpar asitized P. rapae cater pillars, wher e they occurr ed at an av er a ge r elativ e abundance of 32.1% and 19.4%, r espectiv el y.In contr ast, they wer e not or only sporadically detected in parasitized individuals (Fig. 4 ).Conv ersel y, the external microbiome of parasitized P. rapae caterpillars sho w ed some bacterial species that w er e mor e fr equentl y pr esent than others.Specificall y, zOTU3 and zOTU8, both belonging to the genus Pseudomonas , wer e abundantl y pr esent on parasitized individuals, while they were only found at low r elativ e abundances in nonparasitized caterpillars ( < 0.1%) (Fig. 4 ).One bacterial zO TU (zO TU2) was exclusiv el y pr esent in the internal micr obiome of C. glomerata -par asitized cater pillars and absent in any other sample .Moreo ver, it was found in every C. glomeratapar asitized individual anal ysed (Fig. 4 ).This bacterium, identified as Wolbac hia pipientis , occurr ed at an av er a ge r elativ e abundance of 11.2% in C. glomerata -parasitized P. brassicae caterpillars and 48.3% in P. rapae cater pillars par asitized with C. glomerata .Further, zOTU9, identified as Nosema sp., a microsporidium that possesses Ta ble 2. Results of tw o w a y ANOVA on the observed bacterial zO TU ric hness and Shannon div ersity in the inv estigated cater pillars and par asitoid larv ae.Significant differ ences ( P < .05)ar e indicated in bold.a ribosomal unit similar to bacteria (Kawakami et al. 1992 ), was fr equentl y found in parasitized caterpillars.In particular, it was present in all internal samples of C. glomerata -parasitized caterpillars with a r elativ e abundance of 2.7% and 51.6% in P. brassicae and in P. rapae hosts, r espectiv el y.T his zO TU was also found in five of the eight investigated C. rubecula -parasitized P. brassicae individuals (with an av er a ge r elativ e abundance of 0.4%) and in all C. rubecula -parasitized P. rapae individuals (with an av er a ge r elativ e abundance of 64.2%).In contrast, this Nosema species was not detected in any of the nonparasitized caterpillars or in any external samples of the parasitized caterpillars (Fig. 4 ).

Caterpillars
Indicator species analysis confirmed that some bacterial species were specific to some tr eatment gr oups.In particular, for the internal micr obiome, Wolbac hia (zOTU2) and Nosema (zOTU9) were identified as indicators of C. glomerata -parasitized caterpillars of both host species.Nosema (zOTU9) was also highlighted as an indicator of C. rubecula -parasitized P. rapae caterpillars ( Supplementary Table S8 ).Co-occurrence analysis of the external microbiome of P. brassicae caterpillars sho w ed that zOTU4 (Enterobacteriaceae) negativ el y corr elated with ten other zO TUs , suggesting that its pr esence interfer es with the growth of other bac-teria.Similarly, in the external microbiome of P. rapae , the Pseudomonas species corresponding to zOTU8 was negativ el y corr elated with six other species.In the internal microbiome, a strong positiv e co-occurr ence was observ ed between Wolbac hia (zOTU2) and Nosema (zOTU9) in both host species.In contrast, a negative co-occurrence was found between these two species and se v er al zOTUs in P. rapae ( Supplementary Fig. S5 ).Kruskal-Wallis analyses performed on single zOTUs confirmed significant differences in r elativ e abundances between tr eatments for se v er al zO TUs , especially in the internal microbiome of both host species where abundances of zOTU2 ( Wolbachia ) and zOTU9 ( Nosema ) were significantly higher in parasitized than in nonparasitized individuals ( Supplementary Table S9 ).

Taxonomic classification, incidence, and relati v e abundance of par asitoid-larv ae microbes
The same bacteria found abundantly in the internal compartments of parasitized hosts also dominated the microbiomes of par asitoid larv ae (Fig. 4 ; Supplementary Fig. S4 ).P articularl y, the external microbiome of C. glomerata larvae collected from P. brassicae caterpillars was dominated by both Enterococcus (zOTU1) (incidence of 100%; av er a ge r elativ e abundance of 78.4%) and Wolbachia (zOTU2) (100%; 19.4%).When collected from P. rapae , the external microbiome of C. glomerata was particularly dominated by the Wolbachia zOTU, with an av er a ge r elativ e abundance of 94.7% (Fig. 4 ).The internal micr obiome of C. glomerata larv ae was mainly dominated by Wolbac hia , irr espectiv e of the host caterpillar, with a r elativ e abundance of 83.2% in P. brassicae and 94.6% in P .rapae .Additionally , C. glomerata larvae in P. brassicae contained a substantial fraction (13.0%) of Nosema (zOTU9), which was also present in larvae from P. rapae , but at a lo w er average relative abundance (2.2%) (Fig. 4 ).
Similarly, in C. rubecula larvae, a few zOTUs dominated the micr obial comm unities.In the external micr obiome of C. rubecula larv ae collected fr om P. brassicae , Enterococcus (zOTU1) was the most abundant bacterium, with an av er a ge r elativ e abundance of 66.2%.For individuals collected from P. rapae , this Enterococcus zOTU had a r elativ e abundance of 19.4%, and Nosema (zOTU9) and Pseudomonas sp. ( zOTU6) wer e also abundantl y pr esent (Fig. 4 ).The internal microbiome of C. rubecula larvae mainly contained Nosema (zOTU9) and Enterococcus sp.(zOTU1), irr espectiv e of their host, along with se v er al other bacteria that occurred at lo w er relative abundances.In larvae collected from P. brassicae , these zOTUs had a mean r elativ e abundance of 20.0% and 11.1%, r espectiv el y.When P. rapae was the host, the r elativ e abundances were 7.7% for Nosema and 26.7% for Enterococcus (Fig. 4 ).

Bacteria are commonly present in and on host caterpillars and developing parasitoid larvae
Although the effects of parasitism on host micr obial comm unities have been increasingly studied in recent years, particularly in lepidopteran hosts (Cavicchiolli de Oliveira and Consoli 2020 , Gloder et al. 2021, Zhang et al. 2022 ), little is still known about how host micr obial comm unities and those of de v eloping par asitoid larv ae are influenced by both their host and the parasitoid species .Here , through estimation of bacterial abundance by qPCR, we found that bacteria were commonly present in and on the investigated cater pillars, especiall y in P. brassicae , confirming our pr e vious findings (Gloder et al. 2021 ).Furthermor e, high-thr oughput Bacterial taxa represent the most prevalent taxa in the different subgroups based on host caterpillar and parasitism status for caterpillars and host caterpillar and parasitoid species for parasitoid larvae (present at a mean r elativ e abundance > 1% in at least one subgr oup).For eac h zO TU, the a v er a ge r elativ e abundance for eac h subgr oup is giv en in the box as a percenta ge, wher eas the colour indicates pr e v alence (white is absent).zOTUs are identified by a BLAST search against type materials in GenBank.When no significant similarity was found with type materials, the BLAST analysis was performed a gainst entir e GenBank (indicated with and asterisk).Identifications were performed at genus level; when identical scores were obtained for different genera, identifications were performed at family level.When identity percentages were lower than 99%, the percentage of sequence identity with the GenBank entry is given between brackets.Hits with uncultured bacteria are indicated as unidentified bacterium.
Ov er all, cater pillar micr obiomes wer e str ongl y dominated by an Enterococcus species (zOTU1), with an av er a ge r elativ e abundance of up to 97.5% in P. brassicae caterpillars.Although our rarefaction curves tended to r eac h satur ation, pr esumabl y a gr eater sampling depth might still be r equir ed to cover the full diversity of the microbiome in these samples .T he strong dominance of this Enterococcus zO TU ma y ha ve led to under-amplification of other bacterial DN A (May erhofer et al. 2020 ).Although this bacterium was not found in field-collected P. brassicae caterpillars (Gloder et al. 2021 ), this result is consistent with a previous study, where the same Enterococcus zOTU was str ongl y associated with lab-r ear ed P. brassicae caterpillars (Bourne et al. 2023 ).The high relative abundance of this species in lab-r ear ed cater pillars may be linked to the contr olled labor atory conditions under whic h the cater pillars wer e r ear ed and maintained, whic h wer e the same in both studies.Our results also show that the parasitoid larvae collected from the caterpillars possess their own microbiota.Ho w ever, results also sho w ed that the external microbiome of the parasitoid larv ae shar es some similarities with the internal micr obiome of the caterpillar host species, suggesting that there may be an interaction and exchange between the two microbiomes.

Parasitism alters the microbial community composition of host caterpillars: crucial role of host identity
Our r esults clearl y show that par asitism by Cotesia parasitoids significantly alters both the internal and the external microbial com-munity composition of host caterpillars, and that these effects are str ongl y dependent on the host.In a pr e vious study (Gloder et al. 2021 ), parasitism of P. brassicae by C. glomerata altered the internal microbiome of the caterpillars, but no effects were observed on the external micr obiome, possibl y because that study focused on field-collected insects.Differences in microbiomes between natur al and lab-r ear ed insect populations hav e been observ ed fr equentl y, and ar e most pr obabl y due to factors like diet and envir onmental conditions (P ark et al 2019 , Wang et al. 2019, Martínez-Solís et al. 2020 ).We found a strong host-dependent variation in the occurrence of the Enterococcus zO TU (zO TU1).While it remained at high r elativ e abundance on and in parasitized P. brassicae cater pillars, its r elativ e abundance was dr asticall y lo w ered on and in par asitized P. rapae cater pillars compar ed to nonpar asitized cater pillars, irr espectiv e of the par asitoid species.Instead, a higher r elativ e abundance of species belonging to the Pseudomonas genus (zOTU3 and zOTU8) was detected in the external microbiome of parasitized P. rapae individuals, along with a diminished presence of a Serratia species (zOTU5) that was highly abundant on nonparasitized individuals.Some Pseudomonas and Serratia species are known as beneficial bacteria (Teoh et al. 2021, Pons et al. 2022 ), while others may be insect pathogens (Pineda-Castellanos et al. 2015, Flury et al. 2016 ).It is unclear what effect these bacteria had on their host in this study.It has been suggested that C. rubecula is better adapted to P. rapae than to P. brassicae due to differences in host physiology and/or the ability of the parasitoid to regulate these (Harvey et al. 1999 ).Variation in host physiology between P. brassicae and P. rapae may also have favour ed specific micr obes in one host, while adv ersel y affecting them in the other.Further r esearc h is needed to investigate this.

Parasitism alters the microbial community composition of host caterpillars: crucial role of parasitoid identity
In addition to host-dependent variation, our results show that par asitism-induced c hanges in the host micr obiome ar e also determined by the parasitoid species .T his is particularly clear for the internal microbiome of caterpillars parasitized by C. glomerata .Specifically, we found that both P. brassicae and P. rapae cater pillars par asitized with C. glomerata contained a substantial fraction of Wolbachia (zOTU2), which was not detected in nonparasitized caterpillars or in caterpillars parasitized with C. rubecula .Furthermor e, our co-occurr ence anal ysis indicated that this zOTU was negativ el y associated with se v er al zOTUs in P. rapae par asitized cater pillars .T he r elativ e abundance of Wolbac hia was also higher in parasitized P. rapae caterpillars (48.3%) compared to parasitized P. brassicae caterpillars (11.2%).Ho w ever, when comparing the absolute abundance of Wolbachia , estimated by m ultipl ying its r elativ e abundance by the 16S rRNA gene copy number per ul of DNA in each sample, P. brassicae had 1.31 × 10 4 gene copies of Wolbachia per μl of DNA sample, whereas P. rapae had 1.54 × 10 2 gene copies per μl of DNA extract.This suggests that e v en though the r elativ e abundance of Wolbac hia w as lo w in P. brassicae , the bacterium still had a high concentration, higher than in P. rapae , which had a lo w er ov er all bacterial density.In addition, Wolbachia was abundantly found in the developing C. glomerata larvae inside the caterpillar hosts, reaching an average r elativ e abundance of 83.2% and 94.6% in par asitoid larv ae in P. brassicae and P. rapae hosts, r espectiv el y.
Wolbachia is a well-studied genus of intracellular endosymbionts that are commonly found in arthropods .T hese bacteria often manipulate host r epr oduction to favour their own trans-mission (Werren et al. 2008, Sanaei et al. 2020 ) and can benefit their hosts by providing resistance against insecticides and viruses (Berticat et al. 2002, Hedges et al. 2008 ).Wolbachia is estimated to be present in about 80% of lepidopteran species, including species belonging to the Pieridae family (Ahmed et al. 2015a ).Ho w e v er, in our study, Wolbachia was not detected in nonparasitized individuals of P. rapae or P. brassicae , nor in nonparasitized P. brassicae individuals in pr e vious studies (Gloder et al. 2021, Bourne et al. 2023 ).PCR anal ysis using Wolbac hia -specific primers (Doudomis et al. 2012 ) r e v ealed the presence of this bacterium in adult females of our C. glomerata rearing but not in females of C. rubecula , confirming pr e vious r esults (Rattan et al. 2011, Dicke et al. 2020, Gloder et al. 2021 ).Ther efor e, it is reasonable to assume that C. glomerata tr ansferr ed Wolbac hia into the caterpillars during oviposition after which it established and replicated, explaining its high r elativ e abundance in par asitized caterpillars .T his is in line with pr e vious studies showing that parasitoids may transfer Wolbachia into their host during oviposition (Ahmed et al. 2015b ).Alternativ el y, Wolbac hia may be derived from the parasitoid eggs or developing larvae within the host caterpillars, allowing the parasitoid to pass essential symbionts to the next generation, although little is known to support this hypothesis .T he presence of Wolbachia in adult parasitoids could benefit the w asps b y enhancing host-sear ching ability and oviposition frequency (Furihata et al. 2015 ).Howe v er, Wolbac hia may also have negative effects on parasitoids by increasing their susceptibility to hyper par asitoids , i.e .par asitic wasps that attac k the larv ae and pupae of primary parasitoids (van Nouhuys et al. 2016 ).Recent research has suggested that the presence of Wolbachia in parasitized cater pillars c hanges their body odours, pr oviding r eliable cues for hyper par asitoids to locate potential hosts (Bourne et al. 2023 ).Likewise, conspecifics of the primary parasitoid may use these signals to avoid parasitized hosts (Cusumano et al. 2020 ), but further r esearc h is needed to confirm this.While Wolbachia was exclusiv el y associated with cater pillars par asitized with C. glomerata , a Nosema species (zOTU9) was abundantly present within parasitized cater pillars, irr espectiv e of the host or par asitoid species.The species was also abundantly present in developing parasitoid larvae, while it was not found in nonpar asitized cater pillars or the external microbiome of the parasitized caterpillars.Additional PCR analysis using Nosema specific primers (Bosmans et al. 2018 ) on adult females of C. glomerata and C. rubecula from our rearing sho w ed that the Nosema zOTU was also present in se v er al analysed wasps ( Supplementary Fig. S6 ), suggesting that Nosema was tr ansferr ed fr om the par asitoids to the cater pillars during oviposition.This Nosema zOTU was pr obabl y intr oduced in our r earing by r ene wing the par asitoid cultur es with field-collected individuals.Unlike Wolbachia , Nosema is an intracellular microsporidian par asite, r ecentl y r eclassified as a fungus, that is ca pable of infecting a wide range of insects (Yaman et al. 2014, Ia et al. 2017, Bosmans et al. 2018, Galajda et al. 2021 ).Although being an eukaryote, Nosema has a number of prokaryotic features, particularly in its ribosomes (Kawakami et al. 1992 ).A BLAST analysis a gainst GenBank r e v ealed that the two primers used in this study perfectl y matc hed with the small subunit rRNA gene of Nosema , explaining its presence in our data set.The sequence obtained in our study sho w ed a 100% match with Nosema pieriae , a common pathogen in Pieris butterflies (Choi et al. 2002, Yaman et al. 2014 ).The pr olifer ation of this opportunistic pathogen could have been favoured in parasitized individuals as it is known that parasitism causes reduced host immunity responses , which ma y also affect microbial growth (Cavichiolli de Oliveira and Consoli 2020 ).Additionall y, the pr esence of this micr obial par asite might hav e benefitted the de v elopment of the par asitoids b y w eakening their host (Mabbott 2018 ), although further r esearc h is needed to confirm this scenario.
Although the exact mechanisms driving parasitoid-dependent alterations in host microbiomes remain unclear, our data strongly suggest that parasitoid-associated microorganisms can be transferr ed fr om the par asitoid to the cater pillars during oviposition or originate from the developing parasitoid larvae.Many parasitoids release effectors (i.e.molecules that facilitate successful parasitism) into the host that impair the immune system of their hosts.Maternall y tr ansmitted effectors, suc h as symbiotic viruses and v enom, ar e injected during oviposition (Dicke et al. 2020 ).Other effectors, not transmitted by the female parasitoid, include teratocytes (i.e.autonomous cells that detach from the egg membrane during hatching; Strand 2014 ) and secretions released by the par asitoid larv ae (P ang et al. 2023 ).These effectors could, in turn, influence the host microbiome by modulating the host immune system and physiology.Further r esearc h is needed to find out how important they are in shaping the microbiome of host insects.

Conclusions
In summary, our findings demonstrate that endoparasitism by k oinobiont par astoids significantl y affects both the internal and external micr obial comm unities of host cater pillars, and that suc h c hanges depend both on the host and parasitoid species.Our results also show that the developing C. glomerata and C. rubecula larv ae hav e distinct micr obial comm unities .T he internal microbiome of P. brassicae and P. rapae caterpillars parasitized by C. glomerata consistentl y harbour ed Wolbac hia , whic h was entir el y absent in nonparasitized individuals or those parasitized by C. rubecula .Additionall y, par asitized cater pillars sho w ed a high r elativ e abundance of Nosema pieriae , particularly in P. rapae caterpillars.Further inv estigations ar e warr anted to unr av el the potential roles of these microbes in the intricate interactions among the host caterpillar, the parasitoid, and higher tr ophic le v els.

F igure 1 .
Boxplots sho wing the numbers of bacterial 16S rRN A gene copies per μl DN A suspension (logarithmic scale) in the external (A) and internal (B) microbiomes of the investigated caterpillars, as well as in the external (C) and internal (D) microbiomes of the parasitoid larvae collected.Pieris brassicae and P. rapae caterpillars were parasitized with either C. glomerata (CG) or C. rubecula (CR), or remained unparasitized (UN).Samples that were below the detection limit were assigned 2.95 × 10 2 16S rRNA gene copies per μl DNA extr act, whic h corr esponds to the qPCR detection thr eshold.The lo w er and upper whiskers correspond to the minimum and maximum values, with the bar in the middle marking the median value while dots r epr esent outliers.

F igure 3 .
NMDS or dination plots based on Br ay-Curtis distances of r elativ e abundance data of the external and internal micr obiomes of the differ ent caterpillars (A and B) and parasitoid larvae (C and D) samples studied.Pieris brassicae (PB) (circles) and P. rapae (PR) (triangles) caterpillars were parasitized with either C. glomerata (CG) or C. rubecula (CR), or remained unparasitized (UN).Stress values of the plots are 0.165 (A), 0.109 (B), 0.117 (C), and 0.154 (D).

Figure 4 .
Figure 4. Bacterial community profiles of the investigated caterpillars and parasitoid larvae.Pieris brassicae and P. rapae caterpillars were parasitized with either C. glomerata (CG) or C. rubecula (CR), or remained unparasitized (UN).Bacterial taxa represent the most prevalent taxa in the different subgroups based on host caterpillar and parasitism status for caterpillars and host caterpillar and parasitoid species for parasitoid larvae (present at a mean r elativ e abundance > 1% in at least one subgr oup).For eac h zO TU, the a v er a ge r elativ e abundance for eac h subgr oup is giv en in the box as a percenta ge, wher eas the colour indicates pr e v alence (white is absent).zOTUs are identified by a BLAST search against type materials in GenBank.When no significant similarity was found with type materials, the BLAST analysis was performed a gainst entir e GenBank (indicated with and asterisk).Identifications were performed at genus level; when identical scores were obtained for different genera, identifications were performed at family level.When identity percentages were lower than 99%, the percentage of sequence identity with the GenBank entry is given between brackets.Hits with uncultured bacteria are indicated as unidentified bacterium.

Table 1 .
Results of Sc heir er Ray Har e anal ysis on bacterial densities in the external (ext) and internal (int) samples of the investigated caterpillars and parasitoid larvae.Significant differences ( P < .05)are indicated in bold.

Table 3 .
Results of PERMANOVA on the external and internal bacterial community composition of the investigated caterpillars and par asitoid larv ae.Significant differ ences ( P < .05)ar e indicated in bold.